WHOLEMOUNT STAINING AND DECOLORISING OF DROSOPHILA ADULTS
Sean May, 11.08.92. (amended 1.06.93 and 26.11.93)

(Summarised in May.S.T. and O'Kane. C.-.Neurogenetics (1993) 8 (4); 237-238)

Example photomicrographs are available at the same site as this document
labelled   wmount1.gif   and   wmount2.gif  These have been compressed to save 
database size, but still show the efficacy of the procedure.
The line shown was nicknamed 'Gremlin' and no longer exists, it was a GAL4
enhancer-trap line generated by Dr. K. Moffatt in C. O'kanes lab in 
Warwick, it was processed as detailed below.

Dissection

Required:

Dissecting dish (5-10cm tissue culture dishes) containing
approx 5mm depth of polymerised Sylgard (BDH) or similar
transparent silicon rubber.

Fix - 4% paraformaldehyde in PBS (10ml), prepared fresh or
from a frozen 20% stock (paraformaldehyde does not dissolve
readily and can be dissolved by heating (60 C over a long period
 - do not boil), or by alkali addition and re-pH'ing (see Ashburner's
 Drosophila book)

Dissecting pins (as fine as possible - 'minuten pins')

Very sharp forceps (A strongly recommended sharpening stone is
the "original Arkansas"). New forceps are NOT sharp.

Method:

Flies are anaesthetised and dipped in 95% ethanol in order
to reduce their surface tension in aqueous medium, then
transferred into fix in the dissecting dish.
The fly is pinned through the abdomen (ventral side uppermost)
to the Sylgard surface, head towards you.

The front four legs are removed close to their insertion into
the thorax.
Gently tilting the head towards you and extending the
proboscis (if necessary), you will see a transparent skin of
cuticle between the proboscis and neck. break this cuticle by
pinching it SHALLOWLY and pulling. The head will (usually) now
stay tilted towards you.
You will be able to see silvery tracheae in the bottom half of
the head (closest to you), one on each side of the midline,
behind the base of the proboscis, extending for about half of
the visible head capsule.
Insert the forceps and grab the tracheae. The brain is
contained in the other half of the head capsule (away from
you), so you can dig quite deeply without damaging it. Also,
none of the major cerebral nerves are likely to be damaged
from this insertion vantage. Remove all or most of the
tracheae (a pinch followed by a gently tug often removes the
whole air bubble in one go). LEAVE the tubular eye tracheae
(which may be visible) as attempting to remove these can
easily damage the retina.

Your fly should now look like diagram A.

a) On the thorax, two thin, darker lines of cuticle will be
apparent (immediately posterior and lateral to the front leg holes).
Using one pair of forceps to hold one of the lines (deeply,
but as lateral as possible) grab the cuticle lateral to this
point with the second pair and tear sideways (away from the
midline) until the line is broken. Don't remove any cuticle.
Repeat for the other side.
b) Superficially pinch and tear away the cuticle between the
two front leg holes.
c) Similarly, break the lighter cuticle at the same lateral
position on both sides of the thorax as far back as the second
leg hole (Do not work more medially than the outermost edge of
the leg holes.)
d) Grasp the cuticle between the two T2 leg holes (again
superficially) and gently/slowly pull up and towards you until
you can see that it has separated (DO NOT pull too far up as
the ganglia will distort).

Lastly, grasp the cuticle between the T1 leg holes (.......superficially)
and pull up and away from you. The ventral cuticle should be removed
easily to expose the T1 and T2 lobes of the ventral ganglia.
At the most posterior margin of this hole you should see a
tubular trachea underneath where the T2 leg holes were. This
trachae covers the T3 and part of the abdominal neuromere and
can be removed with extreme caution, although it is not
necessary to do this.

Staining:

The carcass should be removed from fix (after a set time) and rinsed
in changes of PBS + 0.1% Triton X-100 for at least five minutes to remove
fix before staining as normal with X-gal. After staining, and
rinsing in PBS at 37 C (to prevent crystal formation) I
recommend a postfix with 4% paraformaldehyde, as I believe
that this reduces subsequent diffusion of the staining during
mounting.
NOTE unfixed tissue may stain more readily, and should be 
the most realistic tissue to look at enhancer trapping/transgenics 
in. The above protocol may therefore be altered to delay the 
fixation to this point, i.e. AFTER the staining (IF the staining is rapid
i.e. minutes/hours, if it takes longer, structural integrity will 
progressively be lost). The dissection can be more difficult in PBS 
(the tissue is more 'sticky') but not much more.

Decolourising:

If the flies have dark eye or cuticle pigment, in order to
visualise internal staining, it is necessary to remove this
pigment. Incubation in a 50/50 mixture of stock hydrogen
peroxide (30%): and 4% paraformaldehyde appears to give the
best result. (i.e. 15% perox; 2% PF) This may be conveniently 
performed at 37 C, Room temperature, or 4 C and the time taken
is variable depending on the pigmentation. 
NOTE the protocol is designed not to remove x-gal staining
as this is already an oxidised product.
CAUTION: Bubbles can often form in the abdominal space, these
can be difficult to remove later and should be removed now if
it is essential to see throughout the abdomen. Bubbles appear to 
form more slowly or not at all at 4 C, although the decolourisation 
takes more time

Mounting:

The carcasses should be transferred through a series of
alcohols to dehydrate them (recommended: 30%, 70%, 2x100% -
going staight to 100% may not dehydrate properly - this will
cause opacity and/or 'silvering' during clearing). They can
then be cleared in Histoclear, Citroclear, or methyl salicylate (oil of
wintergreen )  NOTE Xylene appears to damage the integrity of the x-
gal staining.
The carcasses may then be mounted ventral side up, sandwiched
between fragments of microscope slide to preserve their
orientation (score a glass slide with a diamond pen and break
off rectangles of glass). I recommend Araldite, Durcupan or
similar polymer for permanent preps (EMscope).
Although these polymers will set overnight at 60 C, you run the 
risk of expanded air pockets which will set and 'silver' your preparation. 
Setting at 25 C (or room temp) is highly recommended although
 this can take a week or more. If the carcass is carefully dehydrated 
with mixing and the medium is mixed avoiding bubble formation 
(very tricky to achieve, although MOST of the bubbles can be removed by
centrifuging the mixture in an eppendorf/bench centrifuge) 
then higher setting temperatures can be used.

There is a diagram (dissect.gif) showing the dissection steps, which should be available
from the same source as this text. Similarly, I have a VHS video of a fly brain
dissection that can be sent on arrangement (approx. 5 mins real time dissection
from fly to clean brain) which shows a time efficient procedure.

Sean May (1993). 
lsrei@csv.warwick.ac.uk
ys@dna.bio.warwick.ac.uk